The overall goal of these labeling procedures is to image and quantify stable, dynamic and nascent microtubules in the developing zebrafish embryo. This method can help answer key questions in the field of developmental neurobiology such as, how microtubules contribute to the establishment of cell polarity. The main advantage of this technique is that it optimizes the detection and quantification of distinct microtubule populations in situ.
Generally, individuals new to this method will struggle because of the difficulties in maintaining microtubule integrity. After de-chlorinating staged zebrafish embryos according to the text protocol, transfer them to 1, 5 milliliter centrifuge tubes, removing as much medium as possible. Add one milliliter of 4%PFA in microtubule assembly buffer or MAB, to fix the embryos at 28.5 degree celsius for five minutes.
Then use a pipette to aspirate the fixative and replace it with one milliliter of fresh fixative. Incubate the sampels on a rocker at room temperature for three hours. Aspirate the fixative and add one milliliter of TBS NP-40 buffer.
Gently agitate the embryos on a rocker for five minutes. Repeat the wash two more times. Then, after replacing the buffer with one milliliter of fresh TBS NP-40, store the embryos at four degree celsius for no more than seven days.
After de-yolking and embedding embryos in agarose according to the text protocol, put the sectioning dish filled with TBS NP-40, use a vibratome to generate 40 micrometers sections of the tallest access of the agarose embedded embryos. Use fine forceps to transfer sections of interest to 500 microliters of TBS NP-40 in a 24 well plate. Remove the buffer, and add 500 microliters of blocking solution.
Then rock the sections at room temperature for at least one hour. Replace the blocking solution with 300 microliters of primary antibodies diluted in blocking buffer and incubate the samples on a rocker at four degree celsius for 36 to 72 hours. Use 600 microliters of TBS NP-40 to wash the samples twice.
Placing them on a rocker at room temperature for 30 minutes each. Add 300 microliters of fluorophore conjugated secondary antibodies diluted in blocking buffer to the sections and incubate them in the dark on a rocker at four degree celsius for 16 to 24 hours. Then wash the samples twice as just demonstrated.
Next, add 500 microliters of DAPI solution to the embryos and incubate them at room temperature for 30 minutes. Then use TBS NP-40 to wash the samples three times at room temperature for five minutes. Following the washes, place a drop of mounting medium with anti-fade agent on the center of a dust-free slide.
Use fine forceps to transfer sections to the mounting medium droplet. Then place a dust-free cover slip on top of the sample. Store slides wrapped in foil, in a dry, dark and cool place until imaging is performed.
Using the details outlined in the text protocol, capture z stack confocal images with channel settings for the selected secondary antibody fluorophores and save the image files. With 3D analysis software such as, Image J, open the data file copy. Create a merged image by overlaying the channels of interest by selecting image, color, merge channels.
Select the 594 nanometer, 488 nanometer and DAPI channels to be false colored red, green and blue respectively. Check, create composite and select, okay. Visualize the merged z stack as a single 2D image by performing a maximum intensity projection of the z stack using images, stack, z project.
Enter the starting and ending positions of the inner best z planes as the start slide and stop slice respectively. Select max intensity as the projection type and click okay. In the 3D analysis software, select, create library and provide a descriptive name for the image library.
Click, create and drag raw image files into the library. Next, select a file to analyze. Then, from the view menu, choose, extended focus to display the channel merged image in the main window.
Select, the free hand ROI tool and outline the region of interest to be analyzed. Select the actions tab, followed by, crop to selection to crop the image. Then save the cropped image file under a new name.
Next, click the measurements tab to create the protocol for filtering specific objects relevant to the 3D analysis. Then drag, find objects to the protocol window and rename the first protocol, DAPI. Select the beta tubulin channel in the drop down menu.
Then drag the following settings to the beta tubulin protocol and place them below find objects in the following order. Fill holes and objects, separate touching objects, exclude objects by size and exclude not touching ROIs. Now, select, measure at the bottom of each protocol.
Then for all channels except DAPI, choose intensity and volume measurement and skeletal length for all tubulin labeling. Draw an ROI around the region to be measured. Under the summary tab, observe the measurements after the software processes the region.
Then copy the data and save them to a workable spreadsheet as shown here. To construct a multi-well flow through apparatus, using a jig or a bandsaw, split a 50 milliliter centrifuge tub in half lengthwise. From 70 micrometer nylon mesh, cut seven semicircles with a radius of three centimeters and trim them to fit tightly into one half of the split centrifuge tube.
Using aquarium safe silicon sealer, glue the semicircles into the centrifuge tube parallel to the 10 milliliter gradation markings. After allowing the device to dry for two days, rinse it by soaking it in a beaker of water for two to three hours. Line the top threaded end of the cut centrifuge tube with modeling clay, such that the height of the liquid retained in the flow through device has a depth of one quarter inch.
Prepare the washout apparatus by removing the plunger from a 50 milliliter syringe and insert 12 inches of fine tubing into the tip. Push the tubing in as far as it would go and use modeling clay to seal around the joint. With embryo medium, pre wet the mesh to allow the liquid to run through the entire flow through device.
Angle the device on ice so that liquid pulls in all compartments but still empties out the front where the clay brim is located. Suspend the washout apparatus on a ring stand above the flow through device on ice. Chill 200 milliliters of embryo medium on ice and pour enough into the washout apparatus to ensure that all air bubbles are cleared and that the flow rate is approximately seven milliliters per minute.
Adjust the flow rate by changing the height of the syringe. After de-chlorinating embryos and preparing nocodazole according to the text protocol, use 10 milliliters of cold nocodazole working solution to exchange the embryo medium of the nocodazole treatment group. Place the petri dishes on ice for an appropriate time for the developmental stage.
Then using a fire polished on milliliter glass pipette, transfer the embryos to the flow through apparatus using separate compartments for each experimental group. Start the nocodazole washout by pouring ice cold embryo medium into the top of the 50 milliliter syringe. Allow the microtubules to regrow after 20 minutes of washout at room temperature by using a fire polished one milliliter glass pipette to transfer embryos to glass petri dishes containing warm embryo medium.
As soon as the embryos are transferred, start a timer. Finally, fix the control and washout embryos at one, five and 10 minutes by pipetting approximately 10 embryos into a 1.5 milliliter centrifuge tube filled with one milliliter of 4%PFA MAB fixative and fix the samples as demonstrated earlier in this video. Using glu-tubulin and tyr tubulin as markers for stable and dynamic microtubules.
Dynamic microtubules are shown to pre=dominate in the hind brain at the neural kill stage. As the kill develops into the neural rod, a stage of enhanced epithelialization, qualitatively, fewer microtubules are immunoreactive with the anti tyr-tubulin antibody especially, in the ventral rod. In contrast, glu-tubulin is scattered and punctate throughout the neural kill but is enriched in the ventral neural rod along microtuble tract.
Arrow heads point to specific microtubular bundles or structures where labeling is increased. The nocodazole treatment depolymerizes microtubules resulting in defused labeling. As the microtubules regrow, they extend from the centrosome.
However, this may not be obvious in a single plane due to their non-planar trajectories. Nevertheless, some image analysis software is capable of measuring lengths in 3D, enabling an assessment of microtubule growth after the nocodazole washout. The mean length of microtubules appears to increase over time after the nocodazole washout in all regions of the neural tube analyzed.
Once the data analysis is finished using the 3D image analysis software and quality is controlled for, useful information can be recovered from the raw data such as, average length of total microtubules and stable microtubules with the ratio of stable microtubules to total microtubules. Once mastered, this technique takes no longer than a week from start to finish. While attempting this procedure, it's important to remember to collect and process samples in a timely manner as microtubules depolymerize easily.
This procedure can also be applied to the analysis of mutants. After watching this video, you should have a good understanding of how to capture distinct microtubule populations in the intact embryo.
